Microscopic observation of bacteria: review highlighting the use of environmental SEM
For figures, tables and references we refer the reader to the original paper.
Existing microscopic techniques for bacterial observation
Sampling and bacterial observation are often completed for research purposes. In general, these procedures are not part of the treatment strategy in daily endodontic practice. For bacterial observation related to research, different methods have been adopted throughout the years.
One of the first techniques to observe endodontic pathogens was compound-light microscopy in combination with histological staining and/or sectioning. Although not powerful enough to resolve many structures within the cell, this type of microscope can be used for first stage identification of bacteria by verifying cellular morphology (e.g. rod-, coccal- or spiral-shaped) and the reaction of an organism with the Gram stain (Brown/Brenn staining technique) (Ricucci & Bergenholtz 2003). In addition, markers such as antibodies (monoclonal or specific polyclonal) and nucleic acid probes have been developed for identification at the genus and species level of some bacteria associated with disease as an alternative to lengthy culturing techniques. Additional special equipment (i.e. dark-field illumination and phase-contrast microscopes) has been used to quantify the numbers of motile bacteria in the clinic directly after root canal sampling (Trope et al. 1992). Likewise, in periodontics, some laboratory protocols and a number of chair-side diagnostic tests are routinely used to evaluate large numbers of subgingival plaque samples for their content of a wide range of recognized pathogenic species. Such evaluations are critical in understanding treatment effects and are used by both researchers and clinicians (Greenstein & Polson 1985). The techniques often employed include culturing, dark-field and phase-contrast microscopy (Magnusson et al. 1985, Quirynen et al. 1995), or immunological assays and molecular methods (Socransky et al. 1998, Ximenez-Fyvie et al. 2000, Darby et al. 2001).
A second important approach to bacterial observation used the principle of electron microscopy (EM). The microscope involved constructs an image from a highly focused primary electron (PE) beam, which is scanned over the specimen in a square raster pattern. The PE have a much shorter wavelength than light, and therefore microscopes employing electron beams have 400 times the resolving power of an optical microscope thus revealing much more detail. Employing EM, it was demonstrated that even though bacterial species are diverse in form, their organization is fundamentally similar: small cells about 0.3–10 μm thick, enclosed within a membrane and encased within a rigid cell wall, with no distinct interior compartments. In fact, bacteria, which have a prokaryotic cell structure, do not have membrane-bound organelles within their cells and their DNA is mostly included in a single, closed, circular molecule.
The most common type of electron microscope is called the conventional scanning electron microscope (CSEM), and its associated technique has a long and distinguished record in the field of biomaterials. CSEM offers unique advantages such as high resolution and large depth of field, and related tools have evolved into complex integrated instruments that often incorporate several important accessories. Their principle improvement stems from the method of constructing an image by detecting electron signals generated by the incident beam and emitted from the specimen, whilst scanning across the surface. As a consequence, the whole microscope column, including the sample chamber, operates under high vacuum (<10−5 torr) (1 torr = 133 Pa) to prevent gas scattering of either the incident beam or the produced electrons. The presence of a vacuum, however, implies that samples must not contain any volatile species; they must be solid and dry. Samples that are hydrated in their native state (e.g. biological tissues and cells) must be dried or frozen prior to observation. In addition, this category of samples exhibits low conductivity and has always been a challenge as surface charges generated by the incident electron beam must be drained away to prevent distortion of the image. Coating the specimen with a thin layer of an electrically conductive material will dissipate the electrons and prevent the build-up of charge. However, specimen preparation can introduce artefacts by altering specimen morphology whereas conductive coatings may obscure internal information by impeding the outgoing electron signals (Little et al. 1992). Finally, sample preparation implies that specimens do not preserve their native state. As a result, therapeutic endodontic strategies cannot be observed or tested in situ. The principle of CSEM and its technical issues will be discussed in a further section.
To overcome the limitations of CSEM, a second type of SEM called the environmental scanning electron microscope (ESEM) has been developed. The first commercial version of this product was made by the ElectroScan Corporation (Wilmington, MS, USA; later purchased by FEI/Philips Electron Optics) along with the work of G. D. Danilatos (Danilatos 1988, 1993a) more than a decade ago. In recent years, ESEM has begun to make impact across the diverse field of materials; an expansion which can be evidenced from the increased range of applications over a short span of time (Danilatos 1993b). The major advantage of ESEM is that hydrated and non-conducting samples, such as biological tissues and (bacterial) cells, can be imaged without prior dehydration or conductive coating. ESEM differs therefore from CSEM in two crucial aspects. First, instead of the sample being held under a high vacuum, a gaseous pressure is maintained in the specimen chamber whilst imaging is carried out, although the electron gun itself is kept at standard pressures of around 10−6–10−7 torr. Around the sample, pressures of up to 10–20 torr can be tolerated and all operational parameters can be varied within a range, which is a function of pressure. In this way, if water vapour is the gas in the sample chamber, hydrated samples such as root canal bacteria can be imaged in their ‘native’ state. The second major difference between ESEM and CSEM is that insulators no longer need to be coated with a metallic layer before imaging. Because gas is present in the chamber, a mechanism exists to help dissipate the build-up of charge injected by the incident electron beam. Technically, ESEM is based on the integration of efficient differential pumping with a new design of electron-optics and detection systems. ESEM's physical principles and technical demands will be discussed comprehensively below.
A final type of EM, identified as the transmission electron microscope (TEM), offers unique properties such as high resolution. TEM involves the irradiation of whole specimens or ultra-thin sections (80–90 nm) which are thin enough to transmit at least 50% of the PE (Bancroft & Stevens 1996) using electron beam energies in the range of 60–350 keV. For amorphous materials, contrast is achieved by variations in electron scattering as the electrons traverse the chemical and physical differences within the specimen. The emergent beam of transmitted electrons is focused by a system of lenses to form a magnified, two-dimensional image. The major advantage of TEM is its resolving power. The maximum obtainable resolution (1–2 nm for most biological material) is limited by the nature of the specimen and the techniques involved in specimen preparation. In prevention of artefacts, common preparation procedures of specimens for TEM involve relatively complex and long laboratory processing. Fixed and dehydrated specimens are generally embedded in an epoxy resin and stained with heavy metals (e.g. potassium permanganate or osmium tetroxide) to improve image contrast before ultra-thin sectioning using an ultramicrotome with glass or diamond knives (Nair et al. 2005). For endodontic microbial research, these sections have also been stained with tannic acid and ruthenium red staining prior to examination in the microscope (Haapasalo 1986, Sunde et al. 2002). Other preparation techniques that can be used are cryosectioning and freeze fracturing (followed by freeze etching and the production of a replica) (Haapasalo 1986, Matias et al. 2003). Eventually, the thickness of a section primarily determines the resolution obtainable in a TEM, and therefore the making of sections is very critical in preparing material for fine ultra-structural examination.
Bacterial observation using ESEM opposed to CSEM
CSEM technique and sample preparation
For SEM, basically two types of electron sources can be used to form the electron beam: (i) the thermionic emission (tungsten, Lanthanum Hexaboride or cerium filament) and (ii) the Field-emission (Fe). The latter requires ultra-high vacuum conditions and thus appropriate equipment. The electron beam, which typically has an energy ranging from a few kV to 50 kV, is focused by condenser lenses into a beam with a very fine spot size (∼5 nm). The beam then passes through the objective lens where pairs of scanning coils deflect the beam over a rectangular area of the sample surface. As PE strike the surface they are inelastically scattered by atoms in the ‘spot’ and the beam energy is effectively spread over a certain distance into the sample. Interactions in this region lead to emission of electrons: (i) low-energy secondary electrons (SE or SE-I) (≤50 eV) produced by inelastic collisions with the orbital electrons and dislodged from the specimen itself and (ii) high-energy backscattered (or reflected) electrons (BSE) (>50 eV) that arise from elastic collisions between the PE and the atomic nuclei. These signals of electrons are collected and amplified by a positively biased grid or detector, and results of the analysis are displayed as a specific intensity on screen at a position that represents the position of the incident beam spot. Magnification results from the ratio of the area scanned on the specimen to the area of the screen. Increasing the magnification in a CSEM is therefore achieved quite simply by scanning the electron beam over a smaller area of the specimen. The most common image mode monitors the SE-signal. Because of their low energy these electrons must originate within a few nm (or less) from the specimen surface thus providing topographical information. The brightness of the resulting signal depends on the surface area that is exposed to the primary beam. This area is relatively small for a flat surface but increases for steep surfaces that tend to be brighter; so the final image is pseudo three-dimensional. The emitted SE-signal is detected by a scintillator-photomultiplier device, typically an Everhart–Thornley detector (E–T detector) (Everhart & Thornley 1960). In addition to SE, BSE-signals can also be detected. Because of their much higher energy these electrons may be scattered from fairly deep within the sample resulting in less topographical contrast than the case of SE. BSE have a definite direction. As such, they cannot be collected by a standard SE-detector unless the detector is directly in their path of travel.
Besides the emission and signalling effect, electrons can accumulate on the surface of non-conductive materials and charging will occur. At low beam energies, most SE are generated just below the surface of the specimen and most of them will backscatter into the vacuum. In contrast, high beam energies penetrate much deeper and most of the SE produced cannot escape, thus charging will more likely occur (Egerton et al. 2004). The negative field from the charging surface deflects the incident electron beam from its intended course and causes image drift. Elimination of specimen charging can be achieved by reducing the accelerating voltage below the charging point, or by conductive coating. When the microscope is used at low accelerating voltages, its resolution is greatly reduced. Therefore, the deposition of conductive films is generally preferred.
Apart from their non-conductive properties, biological specimens are, in their native state, hydrated at temperatures above 0 °C. Therefore, such specimens cannot be placed directly into a CSEM as water would evaporate and interfere with the electron generation and detection system thus forming artefacts. In the case of delicate liquid-containing samples, which become hollow when dried, complete collapse often results. To avoid this problem, a complex and extensive series of processing steps is required for reliable observations of hydrated specimens under (ultra) high vacuum. Diverse laboratory protocols have been described in the literature and generally involve successive (pre)fixation, dehydration/drying or freezing, coating with an electron-conductive material and viewing (Watson et al. 1980, Bancroft & Stevens 1996, Van Meerbeek et al. 2000). The main aim of fixation is to preserve the structure of the tissue in an as near life-like condition as possible. Most popular fixative solutions used today are aldehyde fixatives (glutaraldehyde or formaldehyde) made up in phosphate or cacodylate buffers. Dehydration refers to the removal of water from the fixed tissue mass by an organic solvent and is generally carried out in an ascending series of low concentration aqueous ethanol or acetone solutions to an absolute dehydration agent. Once dehydration is complete, the specimen must be dried in a way that causes minimal distortion and disruption of the tissue architecture. Simple air-drying (including evaporation by heat or under vacuum) should be avoided. The most widely used method to prepare biological tissue is known as ‘critical-point drying’ (CPD). This process involves replacement of the absolute ethanol in the tissues with a transitional fluid, most commonly carbon dioxide (CO2). The liquid CO2 is then removed by conversion to its gaseous form by raising the temperature and pressure to the critical point. CPD avoids artefact formation by never allowing a liquid/gas interface to develop; in this way the tissue is not exposed to surface tension forces. An alternative technique called freeze-drying (FD) removes water as vapour directly from ice (sublimation) without passing through the liquid state. FD is so effective in processing chemical components that small living organisms can be preserved and kept alive during and after the process: they pass into a dormant state from which they can be revived by the addition of water (e.g. nutritional supplements industry). In addition to CPD and FD, chemical drying methods, such as HMDS-drying using hexamethyldisilazane have been advocated for biological specimen preparation (Perdigão et al. 1995). Besides specimen dehydration/drying, one can also freeze the specimen (Cryo-SEM) in liquid nitrogen or liquid nitrogen cooled Freon. Freezing prevents the vapour from interfering with the electron generation and detector systems, provided sufficiently low temperatures are reached and maintained during examination. This technique enables a large range of specimens to be successfully examined without dehydration, although it has some limitations including the possibility of ice crystal damage due to a failure to freeze the tissue rapidly enough.